We are indebted to Eleut Hernandez for animal husbandry, Gabe Gaidosh for microscopy expertise, and Khue Tran and Zhenyang Zhao for video editing. This study was funded by National Eye Institute grants R01-EY022129 to JLG and P30 grants EY022589 to UCSD and EY014801 to UM; the American Heart Association, the James and Esther King Foundation, the doctoral student exchange program fund of Fudan University Graduate School (No. 2010033), and an unrestricted grant from Research to Prevent Blindness, Inc.

All animal procedures were approved by the University of California San Diego and University of Miami institutional animal care and use committees (IACUC) and performed in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Visual Research. All reagents and instruments used in surgical procedures are sterile.
1. Anesthetize and Prepare the Rat for Surgery
<ol>
	<li>Prior to procedure, rats are anesthetized with an intraperitoneal injection of ketamine (60 mg/kg) and xylazine (8 mg/kg) according to body weight. Adequate depth of anesthesia should be determined by a negative response to toe pinch stimulus.</li>
	<li>Once anesthetized, pull the tongue forward to prevent asphyxia and apply lubricating ointment to both eyes to prevent drying of the corneas during surgery.</li>
	<li>Shave the surgical sites using a hair clipper and wipe the area three times with 10% providone-iodine detergent solution and 70% ethanol.</li>
	<li>Drape the animal within a sterile field. Sterile gloves and surgical instruments are used during survival surgery. Re-sterilize the tips of instruments using a hot bead sterilizer between animals.</li>
</ol>
2. Surgical Approach
<ol>
	<li>PION induction
	<ol>
		<li>Femoral vein catheterization
		<ol>
			<li>Prepare and clean the surgical site. Shave the right inner thigh using a hair clipper and wipe the area three times each with 10% providone-iodine detergent solution and 70% ethanol.</li>
			<li>Prepare the tubing. Cut a 40cm length of polyethylene tubing (PE 10) sterilized in 70% ethanol. Flush the tubing with saline and connect it to a 1 ml syringe containing a pre-measured solution of 2% erythrosin&#160;B dye (1 &#956;l/mg, yielding a dose of 20 mg/kg body weight). Mount the syringe into a foot-switch controlled infusion pump set to a rate of 600 &#956;l/min.</li>
			<li>Using a No. 15 blade, make a small horizontal incision at the base of the right thigh. Cut and spread the membrane inside and clean the area with sterile cotton swabs.</li>
			<li>Separate the muscle with forceps until the branch of the femoral vein is visible. A sheath surrounds the artery, vein and the nerve. Pinch and pull this sheath upward&#160;with forceps (fine-tip Dumont forceps), and cut a small incision (2-4 mm is usually adequate) near the base of the triangular-shaped wedge with Vannas spring scissors. Expand the cut as necessary.</li>
			<li>Separate the vein and artery with a blunt micro-surgical hook parallel to the direction of the vein. Be careful not to damage the delicate membrane and vein branches. Then, gently lift the vein and separate it from the underlying connective tissue.</li>
			<li>Obtain a needleless nylon suture and place it next to the femoral vein. Using the micro-surgical hook, elevate the vein and pass the&#160;fine-tip forceps beneath the distal region. Grab one end of the suture and pull it underneath the vein. Ligate the distal vein tightly. Pass a second suture in a similar manner under the proximal vein and make a loose knot.</li>
			<li>Make a small cut in the vein near the distal ligation with Vannas spring scissors. Expand the hole as necessary with the fine-tip forceps. Some blood may leak through the cut. Clean the surgical area with cold, sterile BSS and sterile cotton swabs.</li>
			<li>Holding the vein wall at the edge of the cut, catheterize the vessel with the prepared saline-flushed tubing using a needle holder. Tighten the proximal knot around the vein and tubing. Then, anchor the tubing by tying it to the distal suture.</li>
			<li>Check the quality of the catheterization by pressing the foot-switch to inject saline, &#60;1 ml is adequate. Make sure the tubing is unobstructed and has no leaks. Temporarily close the incision with sutures to protect the catheterization and tissue.</li>
		</ol>
		</li>
		<li>Exposure of the optic nerve
		<ol>
			<li>Prepare the surgical site by wiping the pre-shaved area above the left eye three times each with 10% providone-iodine detergent solution and 70% ethanol.</li>
			<li>Make an incision along the skin 2-3 mm behind the eye with a No. 15 blade. Pinch and lift the connective tissue with serrated forceps, and make a small incision with Vannas spring scissors. This small incision is generally about 5 mm in length, but can be longer to provide greater exposure for a&#160;surgeon in training. Continue to bluntly dissect through the connective tissue along the superior rim of the orbital bone, taking care to avoid disrupting blood vessels. Clean the surgical area with cotton swabs.</li>
			<li>Dissect downward through the conjunctiva until the superior rectus muscle is visible. Pinch and dissect through the muscle; the muscle will be liberated from deep within the orbit. Now, surrounding tissues can be utilized to aid in the retraction and elevation of the eye for ease of visualization.</li>
			<li>Retract the flap of skin and connective tissue laterally and downward, and hold in place with a suture and hemostat. This will rotate the eye forward and outward in order to reveal the fat-containing sheath that surrounds the optic nerve.</li>
			<li>Carefully insert a pair of sharp forceps and expand parallel to the optic nerve to separate the connective tissue surrounding the sheath. Do not touch the optic nerve with the sharp tips of the forceps.</li>
			<li>A 5 mm length of the optic nerve and surrounding sheath should now be visible. A network of microvessels on the sheath surface encircles the optic nerve. These will be targeted during laser irradiation.</li>
		</ol>
		</li>
		<li>Intravenous injection of Erythrosin B and laser irradiation
		<ol>
			<li>Wear orange-colored safety glasses at all times while operating the laser irradiation apparatus to shield yourself from the green laser light. Turn on the laser, open the shutter and adjust the peak and average powers of the laser as necessary. Close the shutter.</li>
			<li>Position the rat in the laser irradiation apparatus. To ensure proper beam placement, a weak aiming beam is produced by spatially filtering the laser through a 100 &#181;m diameter hole drilled in the closed shutter blade. Re-expose the optic nerve with a pair of fine tip forceps, and position the aiming beam onto the intraorbital optic nerve between 3 mm and 4 mm behind the optic nerve head.</li>
			<li>Inject the solution of 2% erythrosin&#160;B via activation of the infusion pump. It may circulate for a few seconds while the surgeon adds a small drop of BSS to moisten the surface of the optic nerve.</li>
			<li>Verify the position of the aiming beam and then click the foot-switch to initiate irradiation. An orange-colored safety filter, which is added into the optical path of the microscope, will be immediately triggered followed by the opening of the shutter after a 1 sec delay.</li>
			<li>Irradiate the optic nerve for 90 sec&#160;with a peak power of 135mW and average power of 18mW produced by a beam chopper rotating at 250 Hz with a duty cycle of 15% (this minimizes thermal effects). Yellow fluorescence, visualized as bright orange through the safety filter, will be emitted from the superior surface of the optic nerve and is sufficient to ensure that the beam irradiates&#160;the nerve symmetrically.</li>
			<li>After the irradiation, the orange-colored safety filter will open automatically. Microhemorrhage can be observed in some cases.</li>
			<li>Relieve the traction on the extra ocular muscles and return the eye to a neutral position. Close the incision with interrupted sutures. Then withdraw the catheter&#160;and tie off the femoral vein tightly to prevent leakage; close with interrupted sutures. Apply antibiotic ointment to both incisions. Check the fundus to verify the vascular integrity of the central retinal vein and artery.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Retrograde Labeling of Retinal Ganglion Cells (RGCs). NOTE: In order to evaluate RGC&#160;survival, retrograde labeling with fluorogold (FG) should be completed one week before PION. The method is described in detail in JoVE protocol 81913.
	<ol>
		<li>In brief: anesthetize the animal with ketamine (60 mg/kg) and xylazine (8 mg/kg) and shave the head.</li>
		<li>Surgically scrub the incision site and make a midline incision across the head to expose the skull.</li>
		<li>Drill bilateral holes through the skull (&#248; 2x2 mm) 0.5 mm from both the sagittal and transverse sutures.</li>
		<li>Carefully aspirate the cerebral content that lies over the dorsal surface of the superior colliculus (SC) using a vacuum pump. Then, place a small piece of gelfoam soaked with 4% FG onto the surface of the SC.</li>
		<li>Close the incision with sutures and care for the animal using standard postoperative care.</li>
	</ol>
	</li>
	<li>Post-operative care and Euthanasia
	<ol>
		<li>After surgery, place animal in a separate clean cage on top of a recirculating heated water pad until the animal is recovered.</li>
		<li>Post-surgical analgesics (buprenorphine HCl, 0.01 mg/kg) should be administered twice per day for three consecutive days to minimize discomfort.</li>
		<li>Rat should be kept separately and observed until they are able to maintain sternal recumbency and regain sufficient consciousness.</li>
		<li>Signs of recovery and good health are monitored daily for at least 5 days after the surgery, or until suture removal and adequate healing of the surgical site, whichever occurs latest.</li>
		<li>Euthanize the animals by perfusion with 4% PFA at scientifically appropriate time points after the surgery according to investigative interest.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Wang, Y., Brown, D. P., Duan, Y., Kong, W., Watson, B. D., Goldberg, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+rodent+model+of+posterior+ischemic+optic+neuropathy..">A novel rodent model of posterior ischemic optic neuropathy..</a> JAMA Ophthalmology. 131, 194-204 (2013).</li><li>Rucker, J. C., Biousse, V., Newman, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ischemic+optic+neuropathies.">Ischemic optic neuropathies.</a> Curr Opin Neurol. 17, 27-35 (2004).</li><li>Hayreh, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Posterior+ischaemic+optic+neuropathy:+clinical+features,+pathogenesis,+and+management..">Posterior ischaemic optic neuropathy: clinical features, pathogenesis, and management..</a> Eye (Lond). 18, 1188-1206 (2004).</li><li>Hayreh, S. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inter-Individual+Variation+in+Blood-Supply+of+the+Optic-Nerve+Head..">Inter-Individual Variation in Blood-Supply of the Optic-Nerve Head..</a> Doc Ophthalmol. 59, 217-246 (1985).</li><li>Jacobson, D. M., Vierkant, R. A., Belongia, E. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nonarteritic+anterior+ischemic+optic+neuropathy+-+A+case-control+study+of+potential+risk+factors..">Nonarteritic anterior ischemic optic neuropathy - A case-control study of potential risk factors..</a> Arch Ophthalmol-Chic. 115, 1403-1407 (1997).</li><li>Kosmorsky, G., Straga, J., Knight, C., Dagirmanjian, A., Davis, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+transcranial+Doppler+in+nonarteritic+ischemic+optic+neuropathy..">The role of transcranial Doppler in nonarteritic ischemic optic neuropathy..</a> Am J Ophthalmol. 126, 288-290 (1998).</li><li>Hayreh, S. S., Zimmerman, M. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Non-arteritic+anterior+ischemic+optic+neuropathy:+role+of+systemic+corticosteroid+therapy..">Non-arteritic anterior ischemic optic neuropathy: role of systemic corticosteroid therapy..</a> Graefe's archive for clinical and experimental ophthalmology. 246, 1029-1046 (2008).</li><li>Prado, R., Dietrich, W. D., Watson, B. D., Ginsberg, M. D., Green, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photochemically+Induced+Graded+Spinal-Cord+Infarction+-+Behavioral,+Electrophysiological,+and+Morphological+Correlates..">Photochemically Induced Graded Spinal-Cord Infarction - Behavioral, Electrophysiological, and Morphological Correlates..</a> J Neurosurg. 67, 745-753 (1987).</li><li>Dietrich, W. D., Busto, R., Watson, B. D., Scheinberg, P., Ginsberg, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photochemically+induced+cerebral+infarction.+II.+Edema+and+blood-brain+barrier+disruption..">Photochemically induced cerebral infarction. II. Edema and blood-brain barrier disruption..</a> Acta Neuropathologica. 72, 326-334 (1987).</li><li>Watson, B. D., Dietrich, W. D., Busto, R., Wachtel, M. S., Ginsberg, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Induction+of+reproducible+brain+infarction+by+photochemically+initiated+thrombosis..">Induction of reproducible brain infarction by photochemically initiated thrombosis..</a> Annals of Neurology. 17, 497-504 (1985).</li><li>Watson, B. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal+models+of+photochemically+induced+brain+ischemia+and+stroke..">Animal models of photochemically induced brain ischemia and stroke..</a> Cerebrovascular Disease - Pathophysiology, Diagnosis and Treatment. , 52-73 (1998).</li><li>Watson, B. D., Prado, R., Dietrich, W. D., Ginsberg, M. D., Green, B. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photochemically+induced+spinal+cord+injury+in+the+rat..">Photochemically induced spinal cord injury in the rat..</a> Brain Research. 367, 296-300 (1986).</li><li>Chiu, K., Lau, W. M., Yeung, S. C., Chang, R. C., So, K. 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The authors have nothing to disclose.

Here we describe in detail a method for inducing PION in a rat model. The most critical part of the protocol is the exposure and irradiation of the optic nerve &#8211; to expose the nerve as long as possible while avoiding damage caused by the sharp fine tip forceps or from stretching. In rats, the ophthalmic artery enters the optic nerve &#8804;1 mm from the optic nerve head. Therefore, irradiation of the optic nerve 3-4 mm away from the optic nerve head should only result in ischemia of the capillaries feeding the nerv...

In patients over 50 years old, ischemic optic neuropathy (ION) is the most prevalent type of acute optic neuropathy2. The condition can present as one of two subtypes according to the source of specific affected blood supply&#160;and clinical presentation: anterior (AION) or posterior (PION)3. While the pathogenesis and course of AION have&#160;been studied extensively4-7, PION has remained poorly understood due to its low prevalence, variable presentation, ill-defined diagnostic criteria and lack of an animal model. Furthermore, no treatments have been proven to effectively prevent or reverse vision loss from AION or PION. Therefore, a reproducible and reliable animal model of PION is of great value to study the disease process in vivo and test new therapeutic regimens for neuroprotection and axon regeneration.
Photochemically induced ischemic injury to the microvasculature resulting in vasogenic edema and thrombosis effectively creates regional tissue ischemia8-12. After injection into the vascular circulation, the photosensitive dye erythrosin B produces reactive singlet molecular oxygen upon activation by laser irradiation of&#160;target vessels. The singlet oxygen directly peroxidizes the vascular endothelium, stimulating platelet adherence/aggregation and leading to occlusive thrombus formation. Ischemic damage is spread&#160;to neighboring areas and further exacerbated&#160;by microvascular compression due to vasogenic edema. The overall goal of this protocol is to photochemically induce ischemia in&#160;the retrobulbar optic nerve to mirror the damage caused by PION.
To our knowledge, this is the first model of ischemic injury in the posterior optic nerve1. As this model produces ischemia while avoiding physical trauma, the physiological processes of posterior ischemic optic neuropathy are better mimicked and studied. Also, this model offers a novel platform for screening of candidate therapeutics for optic neuropathies and other CNS ischemic disease. Here, a detailed protocol for femoral vein catheterization, optic nerve exposure, intravenous injection of Erythrosin B and laser irradiation in a rat PION model are described.

<table>
	<tbody>
		<tr>
			<td>532-nm Nd:YAG laser&#160;</td>
			<td>Laser glow</td>
			<td>LRS-532-KM-200-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Beam chopper</td>
			<td>custom-made</td>
			<td>custom-made</td>
			<td></td>
		</tr>
		<tr>
			<td>Mechanical shutter and corresponding shutter drive timer</td>
			<td>AAM Vincent Associates</td>
			<td>SD-10</td>
			<td></td>
		</tr>
		<tr>
			<td>25-cm focal length spherical lens</td>
			<td>CVI/Mellles-Griot</td>
			<td>01 LPX 293</td>
			<td>plano-convexBK7 glass lens with HEBBARTM antireflection coating</td>
		</tr>
		<tr>
			<td>Erythrosin B&#160;</td>
			<td>MP Biomedicals</td>
			<td>190449</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorogold</td>
			<td>Fluorochrome,LLC</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Gelfoam</td>
			<td>Cardinal Health</td>
			<td>CAH1203421</td>
			<td></td>
		</tr>
		<tr>
			<td>Polyethylene tubing (PE10)</td>
			<td>BD Intramedic</td>
			<td>427400</td>
			<td></td>
		</tr>
		<tr>
			<td>No. 10 Blade</td>
			<td>Miltex</td>
			<td>4-110</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Forceps</td>
			<td>F.S.T.</td>
			<td>91150-20</td>
			<td>DUMONT #5 RUSTLESS NON-MAGNETIC</td>
		</tr>
		<tr>
			<td>Forceps with Teeth</td>
			<td>F.S.T.</td>
			<td>11153-10</td>
			<td>Germany stainless</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>F.S.T.</td>
			<td>18025-10&#160;</td>
			<td>Germany stainless</td>
		</tr>
		<tr>
			<td>Vannas spring scissors</td>
			<td>F.S.T.</td>
			<td>2-220&#160;</td>
			<td>JJECK Stainless</td>
		</tr>
		<tr>
			<td>Polyglactin suture</td>
			<td>Ethicon</td>
			<td>J488G</td>
			<td>7-0 suture</td>
		</tr>
		<tr>
			<td>hemostat</td>
			<td>F.S.T.</td>
			<td>12075-12&#160;</td>
			<td>Germany stainless</td>
		</tr>
	</tbody>
</table>

rat model of photochemically-induced posterior ischemic optic neuropathy

Posterior Ischemic optic neuropathy (PION) is a sight-devastating disease in clinical practice. However, its pathogenesis and natural history have&#160;remained poorly understood. Recently, we developed a reliable, reproducible animal model of PION and tested the treatment effect of some neurotrophic factors in this model1. The purpose of this video is to demonstrate our photochemically induced model of posterior ischemic optic neuropathy, and to evaluate its effects with retrograde labeling of retinal ganglion cells. Following surgical exposure of the posterior optic nerve, a photosensitizing dye, erythrosin B, is intravenously injected and a laser beam is focused onto the optic nerve surface. Photochemical interaction of erythrosin B and the laser during irradiation damages the vascular endothelium, prompting microvascular occlusion mediated by platelet thrombosis and edematous compression. The resulting ischemic injury yields a gradual but pronounced retinal ganglion cell dieback, owing to a loss of axonal input - a remote, injury-induced and clinically relevant outcome. Thus, this model provides a novel platform to study the pathophysiologic course of PION,&#160;and can be further optimized for testing therapeutic approaches for optic neuropathies as well as other CNS ischemic diseases.

The resulting ischemic injury induced by this technique yields a gradual but pronounced death of retinal ganglion cells after ischemic axon injury. This is a clinically relevant outcome similar to that observed in the human disease. FG retrograde labeling is used to quantify RGC survival after PION. The same method is employed to validate a successful model creation as well as to assess the effects of&#160;different therapeutic regimens.&#160;Figure 1 shows representative confocal images of FG-positive c...

Watch this Scientific Journal Video about Rat Model of Photochemically-Induced Posterior Ischemic Optic Neuropathy at JoVE.com

Rat Model of Photochemically-Induced Posterior Ischemic Optic Neuropathy

The goal of this protocol is to photochemically induce ischemic injury to the posterior optic nerve in rat. This model is critical to studies of the pathophysiology of posterior ischemic optic neuropathy, and therapeutic approaches for this and other optic neuropathies, as well as of other CNS ischemic diseases.

Posterior Ischemic optic neuropathy (PION) is a sight-devastating disease in clinical practice. However, its pathogenesis and natural history ...

rat-model-photochemically-induced-posterior-ischemic-optic

Research

JoVE Journal

Medicine

8.5K 조회수.  University of Miami Miller School of Medicine.  이 프로토콜의 목적은 화학적 랫트 후부 시신경에 허혈 손상을 유도하는 것이다. 이 모델은 후방 허혈성 시신경 병증의 병태 생리에 대한 연구 및이를위한 치료 적 접근 방법과 다른 광섬유에 중요한 신경 병증,뿐만 아니라 다른 CNS 허혈성 질환이다. 

비디오: Rat Model of Photochemically-Induced Posterior Ischemic Optic Neuropathy

Creation of Reversible Cholestatic Rat Model

Cholestasis is a clinical condition commonly encountered by both surgeons and gastroenterologists. Cholestasis can cause various physiological changes and affect the nutritional status and surgical outcomes. Study of the pathophysiological changes occurring in the liver and other organs is of importance. Various studies have been done in cholestatic rat models.
We used a reversible cholestatic rat model in our recent study looking at the role of methylprednisolone in the ischemia reperfusion injury. Various techniques for creation of a reversible cholestatic model have been described. Creation of a reversible cholestatic rat model can be challenging in view of the smaller size and unique hepatopancreatobiliary anatomy in rats. This video article demonstrates the creation of a reversible cholestatic model.
This model can be used in various studies, such as looking at the changes in nutritional, physiological, pathological, histological and immunological changes in the gastrointestinal tract. This model can also be used to see the effects of cholestasis and various therapeutic interventions on major hepatic surgeries.

Cholestasis is a clinical condition commonly encountered by both surgeons and gastroenterologists. Creation of a reversible cholestatic rat model can be challenging in view of the smaller size and unique hepatopancreatobiliary anatomy in rats. This video article demonstrates the creation of a reversible cholestatic model.

Cholestasis is a clinical condition commonly encountered by both surgeons and gastroenterologists. Cholestasis can cause various physiological changes and ...

Gene Transfer for Ischemic Heart Failure in a Preclinical Model

Various emerging technologies are being developed for patients with heart failure. Well-established preclinical evaluations are necessary to determine their efficacy and safety.
Gene therapy using viral vectors is one of the most promising approaches for treating cardiac diseases. Viral delivery of various different genes by changing the carrier gene has immeasurable therapeutic potential.
In this video, the full process of an animal model of heart failure creation followed by gene transfer is presented using a swine model. First, myocardial infarction is created by occluding the proximal left anterior descending coronary artery. Heart remodeling results in chronic heart failure. Unique to our model is a fairly large scar which truly reflects patients with severe heart failure who require aggressive therapy for positive outcomes. After myocardial infarct creation and development of scar tissue, an intracoronary injection of virus is demonstrated with simultaneous nitroglycerine infusion. Our injection method provides simple and efficient gene transfer with enhanced gene expression. This combination of a myocardial infarct swine model with intracoronary virus delivery has proven to be a consistent and reproducible methodology, which helps not only to test the effect of individual gene, but also compare the efficacy of many genes as therapeutic candidates.

A method of gene transfer for the treatment of ischemic heart failure is described using a swine model of myocardial infarction. Our simple and reproducible method enables us to readily evaluate the efficacy of various gene transfers with a very simple and reproducible way.

Various emerging technologies are being developed for patients with heart failure. Well-established preclinical evaluations are necessary to determine ...

A Simplified Technique for Producing an Ischemic Wound Model

One major obstacle in current diabetic wound research is a lack of an ischemic wound model that can be safely used in diabetic animals. Drugs that work well in non-ischemic wounds may not work in human diabetic wounds because vasculopathy is one major factor that hinders healing of these wounds. We published an article in 2007 describing a rabbit ear ischemic wound model created by a minimally invasive surgical technique. Since then, we have further simplified the procedure for easier operation. On one ear, three small skin incisions were made on the vascular pedicles, 1-2 cm from the ear base. The central artery was ligated and cut along with the nerve. The whole cranial bundle was cut and ligated, leaving only the caudal branch intact. A circumferential subcutaneous tunnel was made through the incisions, to cut subcutaneous tissues, muscles, nerves, and small vessels. The other ear was used as a non-ischemic control. Four wounds were made on the ventral side of each ear. This technique produces 4 ischemic wounds and 4 non-ischemic wounds in one animal for paired comparisons. After surgery, the ischemic ear was cool and cyanotic, and showed reduced movement and a lack of pulse in the ear artery. Skin temperature of the ischemic ear was 1-10 &deg;C lower than that on the normal ear and this difference was maintained for more than one month. Ear tissue high-energy phosphate contents were lower in the ischemic ear than the control ear. Wound healing times were longer in the ischemic ear than in the non-ischemic ear when the same treatment was used. The technique has now been used on more than 80 rabbits in which 23 were diabetic (diabetes time ranging from 2 weeks to 2 years). No single rabbit has developed any surgical complications such as bleeding, infection, or rupture in the skin incisions. The model has many advantages, such as little skin disruption, longer ischemic time, and higher success rate, when compared to many other models. It can be safely used in animals with reduced resistance, and can also be modified to meet different testing requirements.

We have developed a minimally invasive technique to create a rabbit ischemic ear wound model by dividing the central artery and nerve and the cranial neurovascular bundle. A subcutaneous tunnel then cuts all subcutaneous tissues. This procedure causes minimal skin disruption and can be safely used in diabetic animals.

One major obstacle in current diabetic wound research is a lack of an ischemic wound model that can be safely used in diabetic animals. Drugs that work ...

Endothelin-1 Induced Middle Cerebral Artery Occlusion Model for Ischemic Stroke with Laser Doppler Flowmetry Guidance in Rat

Stroke is the number one cause of disability and third leading cause of death in the world, costing an estimated $70 billion in the United States in 20091, 2. Several models of cerebral ischemia have been developed to mimic the human condition of stroke. It has been suggested that up to 80% of all strokes result from ischemic damage in the middle cerebral artery (MCA) area3. In the early 1990s, endothelin-1 (ET-1) 4 was used to induce ischemia by applying it directly adjacent to the surface of the MCA after craniotomy. Later, this model was modified 5 by using a stereotaxic injection of ET-1 adjacent to the MCA to produce focal cerebral ischemia. The main advantages of this model include the ability to perform the procedure quickly, the ability to control artery constriction by altering the dose of ET-1 delivered, no need to manipulate the extracranial vessels supplying blood to the brain as well as gradual reperfusion rates that more closely mimics the reperfusion in humans5-7. On the other hand, the ET-1 model has disadvantages that include the need for a craniotomy, as well as higher variability in stroke volume8. This variability can be reduced with the use of laser Doppler flowmetry (LDF) to verify cerebral ischemia during ET-1 infusion. Factors that affect stroke variability include precision of infusion and the batch of the ET-1 used6. Another important consideration is that although reperfusion is a common occurrence in human stroke, the duration of occlusion for ET-1 induced MCAO may not closely mimic that of human stroke where many patients have partial reperfusion over a period of hours to days following occlusion9, 10. This protocol will describe in detail the ET-1 induced MCAO model for ischemic stroke in rats. It will also draw attention to special considerations and potential drawbacks throughout the procedure.

Several animal models of cerebral ischemia have been developed to simulate the human condition of stroke. This protocol describes the endothelin-1 (ET-1) induced middle cerebral artery occlusion (MCAO) model for ischemic stroke in rats. In addition, important considerations, advantages, and shortcomings of this model are discussed.

Stroke is the number one cause of disability and third leading cause of death in the world, costing an estimated $70 billion in the United States in ...

Angiogenesis in the Ischemic Rat Lung

The adult lung is perfused by both the systemic bronchial artery and the entire venous return flowing through the pulmonary arteries. In most lung pathologies, it is the smaller systemic vasculature that responds to a need for enhanced lung perfusion and shows robust neovascularization. Pulmonary vascular ischemia induced by pulmonary artery obstruction has been shown to result in rapid systemic arterial angiogenesis in man as well as in several animal models. Although the histologic assessment of the time course of bronchial artery proliferation in rats was carefully described by Weibel 1, mechanisms responsible for this organized growth of new vessels are not clear. We provide surgical details of inducing left pulmonary artery ischemia in the rat that leads to bronchial neovascularization. Quantification of the extent of angiogenesis presents an additional challenge due to the presence of the two vascular beds within the lung. Methods to determine functional angiogenesis based on labeled microsphere injections are provided.

The lung is perfused by both the systemic bronchial artery and pulmonary arteries. In most lung pathologies, it is the smaller systemic vasculature that shows robust neovascularization. Cessation of pulmonary blood flow promotes brisk bronchial angiogenesis. We provide surgical details of inducing left pulmonary artery ischemia that promotes bronchial neovascularization.

The  adult lung is perfused by both the systemic bronchial artery and the entire  venous return flowing through the pulmonary arteries. In most lung ...

A Piglet Model of Neonatal Hypoxic-Ischemic Encephalopathy

Birth asphyxia, which causes hypoxic-ischemic encephalopathy (HIE), accounts for 0.66 million deaths worldwide each year, about a quarter of the world&#8217;s 2.9 million neonatal deaths. Animal models of HIE have contributed to the understanding of the pathophysiology in HIE, and have highlighted the dynamic process that occur in brain injury due to perinatal asphyxia. Thus, animal studies have suggested a time-window for post-insult treatment strategies. Hypothermia has been tested as a treatment for HIE in pdiglet models and subsequently proven effective in clinical trials. Variations of the model have been applied in the study of adjunctive neuroprotective methods and piglet studies of xenon and melatonin have led to clinical phase I and II trials1,2. The piglet HIE model is further used for neonatal resuscitation- and hemodynamic studies as well as in investigations of cerebral hypoxia on a cellular level. However, it is a technically challenging model and variations in the protocol may result in either too mild or too severe brain injury. In this article, we demonstrate the technical procedures necessary for establishing a stable piglet model of neonatal HIE. First, the newborn piglet (&#60; 24 hr old, median weight 1500 g) is anesthetized, intubated, and monitored in a setup comparable to that found in a neonatal intensive care unit. Global hypoxia-ischemia is induced by lowering the inspiratory oxygen fraction to achieve global hypoxia, ischemia through hypotension and a flat trace amplitude integrated EEG (aEEG) indicative of cerebral hypoxia. Survival is promoted by adjusting oxygenation according to the aEEG response and blood pressure. Brain injury is quantified by histopathology and magnetic resonance imaging after 72 hr.

Hypoxic-ischemic encephalopathy following perinatal asphyxia can be studied using animal models. We demonstrate the procedures necessary for establishing a piglet model of neonatal hypoxic-ischemic encephalopathy.

Birth asphyxia, which causes hypoxic-ischemic encephalopathy (HIE), accounts for 0.66 million deaths worldwide each year, about a quarter of the ...

Demonstration of the Rat Ischemic Skin Wound Model

The propensity for chronic wounds in humans increases with ageing, disease conditions such as diabetes and impaired cardiovascular function, and unrelieved pressure due to immobility. Animal models have been developed that attempt to mimic these conditions for the purpose of furthering our understanding of the complexity of chronic wounds. The model described herein is a rat ischemic skin flap model that permits a prolonged reduction of blood flow resulting in wounds that become ischemic and resemble a chronic wound phenotype (reduced vascularization, increased inflammation and delayed wound closure). It consists of a bipedicled dorsal flap with 2 ischemic wounds placed centrally and 2 non-ischemic wounds lateral to the flap as controls. A novel addition to this ischemic skin flap model is the placement of a silicone sheet beneath the flap that functions as a barrier and a splint to prevent revascularization and reduce contraction as the wounds heal. Despite the debate of using rats for wound healing studies due to their quite distinct anatomic and physiologic differences compared to humans (i.e., the presence of a panniculus carnosus muscle, short life-span, increased number of hair follicles, and their ability to heal infected wounds) the modifications employed in this model make it a valuable alternative to previously developed ischemic skin flap models.

The rat, due to its size, availability, and rather docile behavior, has been utilized as a research model for many years. The goal of this protocol is to utilize the rat as an ischemic skin wound healing model to provide valuable insight into the pathophysiology of chronic wounds.

The propensity for chronic wounds in humans increases with ageing, disease conditions such as diabetes and impaired cardiovascular function, and ...

The Rodent Model of Nonarteritic Anterior Ischemic Optic Neuropathy (rNAION)

Nonarteritic anterior ischemic optic neuropathy (NAION) is a focal ischemic lesion of the optic nerve that affects 1/700 individuals throughout their lifetime. NAION results in optic nerve edema, selective loss of the retinal ganglion cell neurons (RGCs) and atrophy of the optic nerve. A rodent model of NAION that expresses most NAION features and sequelae has been developed, which is applicable to both rats and mice. This model utilizes a focal laser application of 532 nm wavelength to illuminate a photoactive dye, Rose Bengal (RB), to cause capillary damage and leakage at the targeted anterior optic nerve (the laminar region). After rNAION induction, there is an early optic nerve ischemia, optic nerve edema, and intraneural inflammation, followed by selective RGC and axonal loss. Since the optic nerve is a CNS white matter tract, the rNAION model is applicable to mechanistic studies of selective white matter ischemia, as well as neuroprotective analyses and short and long-term mechanisms of glial and neuronal response to ischemia.

The Rodent Model of Nonarteritic Anterior Ischemic Optic Neuropathy rNAION

The following report describes how to replicate the rodent model of nonarteritic anterior ischemic optic neuropathy (rNAION), using the appropriate dye, contact lens and laser parameters. We also reveal the appropriate steps for evaluating the rNAION lesion in vivo.

Nonarteritic anterior ischemic optic neuropathy (NAION) is a focal ischemic lesion of the optic nerve that affects 1/700 individuals throughout their ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

Oxygen-Induced Retinopathy Model for Ischemic Retinal Diseases in Rodents

One of the commonly used models for ischemic retinopathies is the oxygen-induced retinopathy (OIR) model. Here we describe detailed protocols for the OIR model induction and its readouts in both mice and rats. Retinal neovascularization is induced in OIR by exposing rodent pups either to hyperoxia (mice) or alternating levels of hyperoxia and hypoxia (rats). The primary readouts of these models are the size of neovascular (NV) and avascular (AVA) areas in the retina. This preclinical in vivo model can be used to evaluate the efficacy of potential anti-angiogenic drugs or to address the role of specific genes in the retinal angiogenesis by using genetically manipulated animals. The model has some strain and vendor specific variation in the OIR induction which should be taken into consideration when designing the experiments.

Oxygen-induced retinopathy (OIR) can be used to model ischemic retinal diseases such as retinopathy of prematurity and proliferative diabetic retinopathy and to serve as a model for proof-of-concept studies in evaluating antiangiogenic drugs for neovascular diseases. OIR induces robust and reproducible neovascularization in the retina that can be quantified.

One of the commonly used models for ischemic retinopathies is the oxygen-induced retinopathy (OIR) model. Here we describe detailed protocols for the OIR ...